Fabrication and characterisation of bioartificial sponges for auricular cartilage engineering

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Abstract

Auricle reconstruction due to congenital, post-infective or post-traumatic defects represents one of the most challenging procedure in the field of aesthetic and reconstructive surgery due to the highly complex three-dimensional anatomy of the external ear. In the last two decades tissue engineering strategies are under investigation as potential alternatives to overcome the shortcomings related to the harvesting of autogenous rib cartilage grafts employed in current gold standard surgical reconstruction procedure. In the present study, Poly(vinyl alcohol)/gelatin (PVA/G) sponges at different weight ratios [PVA/G 90/10, 80/20, 70/30 (w/w)%] were produced via emulsion and freeze-drying and finally cross-linked by exposure to glutaraldehyde vapors. Since the scaffolds aim to act as a preliminary extracellular matrix, the addition of a polysaccharide (i.e. alginate) was considered in this study [PVA/G/Alg 80/10/10, 90/5/5 (w/w)%]. However, the PVA/G/Alg foam did not maintain a suitable shape after freeze-drying. Differently, PVA/G sponges gave rise to highly porous, water stable and hydrophilic scaffolds, which were selected for further characterisation. Morphological, physico-chemical and biomechanical characterisation of PVA/G sponges showed interesting properties potentially useful for cartilage regeneration, such as round-shaped interconnected pores, high swelling capacity due to PVA and G interaction and chemical cross-linking, and essentially elastic behaviour. The PVA/G sponge 70/30 (w/w)% was selected since it presented a higher gelatin content and no significant differences in morphological, physico-chemical and mechanical properties when compared to PVA/G 80/20 (w/w)%. Biological studies were performed using the in vitro cell culture method. Since chondrocytes are difficult to isolate and expand in large numbers, bone marrow derived human mesenchymal stromal cells (hMSCs) were used, to be differentiated into chondrocytes. Different culture conditions were tested to optimise hMSC chondrogenesis on these scaffolds: commercial versus hand-made differentiation medium, undifferentiated versus pre-differentiated hMSC seeding, and static culture versus low-intensity ultrasound (US) stimulation. Cell viability throughout culture time proved that PVA/G is a suitable scaffold for hMSC adhesion, growth and chondrogenic differentiation. Experimental results highlighted an intense glycosaminoglycan (GAG), glycoprotein and collagen synthesis after three weeks of differentiation in a commercially available differentiative medium. Immunohistochemistry for chondrogenic markers revealed an early differentiation stage, characterised by the expression of Sox-9, a chondrogenic transcription factor, and type I collagen fibers. An initial expression of aggrecan was observed, while type II collagen and elastin were not detected at protein level. The application of US on cell/scaffold constructs proved an enhanced chondrogenic differentiation in terms of extracellular matrix deposition and a 30% higher type II collagen gene expression than those observed in the non-US stimulated counterparts. This study demonstrated that 70/30 PVA/G sponge is a suitable candidate for auricle reconstruction. Low-frequency US stimulation could represent a valuable tool to further improve chondrogenic differentation.

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List of Figures

Figure 1 – Schematic illustration of MSC differentiation stages...17 Figure 2 - SEM analysis: A) view of PVA/G 70/30 (w/w)% surface; B) view of PVA/G 80/20 (w/w)% surface; C) PVA/G 90/10 (w/w)% surface; D) pore diameter distribution at different (w/w)%...33 Figure 3 – Volume swelling ratio of PVA/G blends at different g concentrations (90/10, 80/20, 70/30 w/w%)...34 Figure 4 - DSC thermograms of crosslinked and uncrosslinked PVA/G samples at different gelatin concentrations...35 Figure 5 - FTIR spectra of crosslinked and uncrosslinked PVA/G samples at different gelatin concentrations...36 Figure 6 - Stress-time experimental data at different strain rates within the LVR ...38 Figure 7 – Histochemistry, immunohistochemistry and SEM analysis on chondroinduced hMSCs cultured on a PVA/G scaffold...39 Figure 8 - Viability of chondroinduced hMSCs over differentiation time...40 Figure 9 – Histochemistry and SEM analysis on chondroinduced hMSCs cultured on PVA/G scaffolds. First column: Hematoxylin-Eosin staining; second column: SEM images...42 Figure 10 – Histochemistry on chondroinduced hMSCs cultured on PVA/G scaffolds. First column: PAS staining; second column: Alcian Blue pH 2.5 staining...43 Figure 11 – Histochemistry on chondroinduced hMSCs cultured on PVA/G scaffolds. First column: Alcian Blue pH 1 staining; second column: Toluidine Blue staining...44 Figure 12 – Histochemistry on chondroinduced hMSCs cultured on PVA/G scaffolds. First column: Orcein-Van Gieson staining; second column: Alcian Blue pH 1-Orcein-Van Gieson staining...45 Figure 13 – Immunohistochemistry on hMSCs cultured on PVA/G scaffolds. First column: Sox-9; second column: aggrecan...46

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6 Figure 14 – Immunohistochemistry on hMSCs cultured on PVA/G scaffolds. First column: type I collagen; second column: type II collagen ...47 Figure 15 – Histochemistry and SEM analysis on pre-differentiated chondroinduced hMSCs cultured on PVA/G scaffolds. First row: Hematoxylin-Eosin staining; second row: SEM images; third row: PAS staining; fourth row: Alcian Blue pH 2.5 staining...49 Figure 16 – Histochemistry on pre-differentiated chondroinduced hMSCs cultured on PVA/G scaffolds. First row: Alcian Blue pH 1 staining; second row: Toluidine Blue staining; third row: Orcein-Van Gieson staining; fourth row: Alcian Blue pH 1-Van Gieson staining...50 Figure 17 – Immunohistochemistry on pre-differentiated chondroinduced hMSCs cultured on PVA/G scaffolds. First row: sox-9; second row: aggrecan; third row: type I collagen; fourth row: type II collagen...51 Figure 18 – Beta-actin and type II collagen gene expression evaluated by RT-PCR...66 Figure 19 – Relative fold levels of type II collagen gene expression normalised to beta-actin utilised as loading control...66 Figure 20 – Immunohistochemistry on chondroinduced hMSCs cultured on a PVA/G scaffold in dynamic culture conditions...68

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List of Tables

Table 1 - Phase transition temperatures of PVA/G sponges at different (w/w)% ratios before and after cross-linking with glutaraldehyde...35 Table 2 - Physical significance of PVA/G sponges absorption peaks and corresponding incident radiation wavenumber...36 Table 3 - Apparent compressive elastic modulus of PVA/G sponges at different strain rates...37 Table 4 - PVA/G viscoelastic parameters derived from fitting of experimental data...38

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Abbreviations

AM - Additive Manufacturing ANOVA - Analysis of variance ASCs - Adipose-derived Stem Cells BSA - Bovine Serum Albumin CAD - Computer-aided-design

CSPCs - Cartilage Stem/Progenitor cells CT - Computed Tomography

DMEM/F-12 - Dulbecco’s Modified Eagle’s Medium/ Nutrient Mixture F-12 DMEM-LG - Dulbecco’s Modified Eagle’s Medium-Low Glucose

DPX - Distyrene Plasticizer Xylene DSC - Differential Scanning Calorimetry ECM - Extracellular Matrix

FBS - Fetal Bovine Serum

FTIR - Fourier Transformed Infrared spectroscopy G - Gelatin

GAGs - Glycosaminoglycans GTA - Glutaraldehyde

HDPP - High-Density Porous Polyethylene hMSCs - human Mesenchymal Stromal Cells IgG - Immunoglobulin G

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9 ITS - Insulin-Transferrin-Selenium

LIUS - Low-intensity Ultrasounds PAS - Periodic Acid-Schiff PBS - Phosphate Buffered Saline PCR – Polymerase Chain Reaction PGA - Poly(glycolic acid)

PLA - Poly(lactic acid) PVA - Poly(vinyl alcohol) SDS - Sodium Dodecyl Sulfate SEM - Scanning Electron Microscopy SLS - Standard Linear Solid

SOX9 - Sex Determining Region box 9 TGF-β1 - Transforming Growth Factor-beta 1 US – Ultrasounds

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Chapter 1 Outline

This document is divided into six chapters. Firstly, an overview of the clinical problem and tissue engineering approaches adopted in cartilage regeneration are presented in Chapter 2-Introduction, as well as the aim of the present thesis. Chapter 3-Materials and Methods describes in detail the experimental design and materials and methodology employed to perform the study. All relevant results are reported in Chapter 4- Results and discussed in the following Chapter 5-Discussion. Finally, the main results are highlighted and suggestions for future work are presented in Chapter 6-Conclusions.

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Chapter 2 Introduction

2.1 Clinical problem: malformation or absence of the auricle

Since 1940s external ear surgical reconstruction to correct auricle loss due to congenital, post-infective or post-traumatic defects represents one of the most challenging procedure in the field of aesthetic and reconstructive surgery (Conway H., 1948). What makes this surgical task particularly challenging is the highly complex three-dimensional anatomy of the external ear (Kamil S.H., 2004; Han S.E., 2015). Microtia and anotia are the most frequent congenital defects that can cause abnormalities or absence of auricular cartilage. The term microtia, Greek for ‘little ear’, refers to a spectrum of congenital malformations of the auricle which ranges from mild structural abnormality to the complete absence of the external ear, denoted as anotia (Griffin M.F., 2016). Microtia-anotia can occur as an isolate condition or as consequence of a congenital syndrome which affects craniofacial development, such as Hemifacial microsomia or Treacher Collins syndrome (Luquetti D.V., 2011; Griffin M.F., 2016). Even though microtia aetiology is not clear, environmental factors, particularly altitude, and exposure to retinoids, alcohol, thalidomide and mycophenolate during pregnancy appears to influence microtia incidence (Luquetti D.V, 2011). The estimated prevalence is between 0.8 and 17.4 per 10000 live births in different geographical areas: higher prevalence rates are recorded in Ecuador, Cile e Finlandia with prevalence values of 17.4, 8.8 e 4.3 per 10000 births respectively (Luquetti D.V., 2011). As highlighted by epidemiological studies, microtia-anotia is unilateral in 79-93% of cases and right ear is most frequently affected, with higher incidence rate in males(Luquetti D.V., 2011). Language skills are not compromised by unilateral microtia-anotia, since there is normal hearing in the healthy ear. However, children affected by this malformation are exposed to a higher risk of delays in language development and attention deficit disorders (Luquetti D.V., 2011). Further studies underline that deformity has a negative effect on self-confidence, life quality and psychosocial development of the child (Zhou G., 2018). The aforementioned problems related to auricle abnormality or absence demonstrate the impact of auricle reconstruction on patients’ life.

2.2 Surgical auricle reconstruction

The main steps in the history of surgical auricle reconstruction could be represented by:

 Sixteenth century: first attempts to repair partial losses of external ear by using skin flaps (Conway H., 1948);

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12  1935: Esser was the first to employ an allomaterial to improve reconstructed ear mechanical

stability (Conway H., 1948);

 1940: the first cadaver cartilage graft was implanted to reconstruct patient’s auricle (Conway H., 1948);

 1947: Brown asserted that autologous costal cartilage can be used in auricle reconstruction (Conway H., 1948);

 2001: Thorne et al suggested the employment of a prosthetic auricle as an alternative to surgical reconstruction. The application of the prosthesis, directly upon skin by using adhesive tissue or by fixing it to a small metallic button implanted in the temporal bone, provides good aesthetic results minimizing risks associated with surgical reconstruction (Thorne C.H., 2001). However difficulty of retention, skin irritation and insufficient compliance limit the potential benefits of an aesthetic prosthesis implantation (Thorne C.H., 2001).

 Nowadays: current surgical treatments include the use of rib cartilage autograft and alloplastic biomaterials for auricle reconstruction (Kamil S.H., 2004; Jessop Z.M., 2016; Griffin M.F., 2016).

2.2.1 State of art

The procedure of microtia reconstruction generally involves three surgical steps: the implantation of a tissue expander, pinna reconstruction by using an autologous cartilage graft, the reconstruction of tragus and concha (Jiang H., 2008). Despite decades of experience, total surgical reconstruction of the external ear still remains one of the biggest challenges in aesthetic surgery (Ciorba A., 2006). Multiple factors affect the success of the reconstruction. First of all an adequate amount of healthy tissue and good blood circulation in auricular region are required (Han S.E., 2015). Moreover, the reconstructed ear should have same size, shape, height and color of the healthy ear, mechanical stiffness similar to that of native auricle, correct cephalic-auricular angle and ability to retain shape and dimension after the implantation in order to guarantee the acceptance of the result (Han S.E., 2015). In addition to these considerations, morbidity, inevitable lesion at the donor site, risk of pneumothorax, postoperative pain and visible chest-wall deformity associated with rib cartilage harvesting must be taken into account (Ciorba A., 2006; Griffin M.F., 2016; Jessop Z.M., 2016). Rib cartilage shows a fibrocartilegineous nature and thus it is inherently different in terms of flexibility and mechanical strength from elastic auricular cartilage and susceptible to progressive calcification (Han S.E., 2015; Jessop Z.M., 2016). These shortcomings related to surgical reconstruction procedure highlighted the need of an alternative to the use of autogenous rib cartilage grafts which currently represents the gold standard in auricle reconstruction. In the last two decades tissue engineering

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13 is under investigation as an alternative to obtain ear-shaped cartilagineous implants. Tissue engineering approaches will be discussed in more detail in section 2.5.

2.3 Ear anatomy

Ear is anatomically divided in three regions: external ear, middle ear and inner ear. The external ear has an aesthetical significance and serves as a reverberation chamber (Zopf D.A., 2015); middle ear is responsible for amplification and transmission of sound waves to appropriate regions of the inner ear; inner ear is the location of balance and hearing sense organs. The external ear consists of the auricle (or pinna) and the ear canal. The auricle represents the visible part of the external ear and it is composed of elastic cartilage. The role of the auricle is to protect the ear canal (or external acoustic meatus), preventing the entrance of foreign bodies, and to channel sound waves into the tympanic membrane (Zopf D.A., 2015). The tympanic membrane is a thin membrane of connective tissue located at the border between external and middle ear. The tympanic cavity, separated from the external meatus by the tympanic membrane, is in communication with the nasopharynx through the auditory tube. The auditory tube (or Eustachian tube) is a 4 cm conduit within the temporal bone. The end connected to the tympanic cavity is relatively narrow and supported by elastic cartilage, while the outlet in the nasopharynx is larger and has funnel-like shape. Functionally the auditory tube allows to equilibrate the pressure in the tympanic cavity with atmospheric pressure, thus avoiding tympanic membrane distortion. The tympanic cavity contains the auditory ossicles: malleus, incus and stapes. These bones are among the smallest in the human body. The lateral portion of the malleus is connected with the inner surface of the tympanic membrane in three points; the incus connects the malleus with the stapes; the base of the stapes is attached to the membrane of the oval window, an opening in the wall of middle ear cavity. Sound waves reach the tympanic membrane, causing the vibration of the membrane at sound frequency, between 20 and 20000 Hz. The vibrations of the tympanic membrane are converted into auditory ossicles chain movements which in turn generate pressure changes in the perilymph present in the cavity of the inner ear. Two muscles, tensor tympani and stapedius, limit the extent of the tympanic membrane and ossicles movements in case of intense noise exposure. The centre of balance and hearing senses is located in the inner ear. The inner ear consist of a membranous labyrinth and a bony labyrinth. The membranous labyrinth is a complex system of communicating channels and chambers containing a liquid, called endolymph. The bony labyrinth encloses and protects the membranous labyrinth and is externally fused with the temporal bone. A liquid, called perilymph, having properties similar to cerebrospinal fluid, fills the space between membranous and bony labyrinth. The bony labyrinth can be divided in three parts: the vestibule, the semicircular canals and the cochlea. The vestibular system consists of the vestibule and the semicircular canals. Receptors responsible for balance and spatial orientation are located in the walls of the vestibular system, while sensory receptors of hearing are located within the cochlea. The cochlea consists of a narrow and elongated portion of the membranous

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14 labyrinth, included between two perilymphatic chambers wrapped in a spiral-shape around a small conical bone, called modiolus. The receptors of the inner ear are represented by specialized epithelial cells, hair cells, characterised by the presence of long stereocilia at the apical pole. The hair cells are mechanoreceptors senisitive to stereocilia distortion due to pressure changes in the perilymph, forming the organ of Corti. The area of maximum stimulation of the organ of Corti varies according to the frequency of the sound, while the magnitude of the distortion allows to detect sound intensity. Auditory cells stimulation activates sensory neurons responsible for sound perception (Martini F.H., 2015).

2.4 Auricular cartilage properties and composition

The auricle, the external auditory meatus and the Eustachian tube consists of elastic cartilage (Bloom W., 1968; Ciorba A., 2006). The properties of auricular cartilage depend on characteristics and spatial distribution of its main structural components: collagen fibers, elastic fibers, glycosaminoglycans (GAGs), chondrocytes and water (van Osch G.J., 1998). Chondrocytes are roundly-shaped and can be observed as single cells or grouped in isogenic groups composed by two or three cells surrounded by abundant extracellular matrix (ECM) (Bloom W., 1968; Adamo S., 2012). Collagen fibers, essentially composed of type II collagen, are responsible for tensile strength, while elasticity and flexibility of the tissue are due to the presence of elastic fibers, peculiar to elastic cartilage (Gosline J., 2002). Elastic fibers are obtained by polimerization of tropoelastin molecules synthesized by cells (Bloom W., 1968; Adamo S., 2012). Elastic fibers also contains a microfibrillar component consisting of fibrillin, which plays a structural role by guiding tropoelastin orientation (Bloom W., 1968; Adamo S., 2012). GAGs contain negatively charged sulfate groups able to attract water molecules and bind proteoglycans to collagen fibrils, providing unique biomechanical properties (Ross M.H., 2016). Several GAGs covalently attached to a core protein form a proteoglycan. Cartilage predominant proteoglycan is aggrecan, which can be considered the principle responsible for cartilage elasticity and viscosity retention (Xia P., 2017). ECM of elastic cartilage also contains smaller proteoglycans such as biglycan, decorin and fibromodulin, glycoproteins as fibronectin and other non-collagenous proteins (van Osch G.J., 1998).

2.4.1 Biomechanical characteristics of elastic cartilage

Auricular cartilage, as any other biological tissue, shows a viscoelastic behaviour. Elastin behaves biomechanically as a linear elastic solid and can be elestically deformed up to 160% strain values (Fung, 1997). The stress-strain curve obtained by mechanical testing of elastin samples is essentially linear, even though there is a slight difference between loading and unloading curve due to a phenomenon of energy dissipation within the material (Fung, 1997). Elastin elasticity is a direct consequence of the entropic recoil of elastin molecules (Urry D.W., 2002). Collagen is a basilar structural protein able to provide mechanical

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15 integrity and strength to soft tissues, thus playing a key role in determining tissue biomechanical properties (Fung, 1997).

Data dealing with auricular cartilage mechanical properties available in literature are limited. Griffin et al tested samples of human auricular cartilage harvested from cadavers (average age 69±10) in order to explore the biomechanical properties of this tissue (Griffin M.F., 2016). Static compressive test reported an overall elastic modulus of 1.66±0.63 MPa without significant differences among the different anatomical regions in which human auricle can be divided (Griffin M.F., 2016). Elastic cartilage does not undergo calcification but its biomechanical properties change as a consequence of ageing (Bloom W., 1968; Adamo S., 2012). Individual’s age advancing is related to significant changes in matrix thickness and composition: the amount of elastin decreases, elastic fibers are thinner and more fragmented, chondrocytes number and size decrease (Nimeskern L.U., 2016; Riedler K.L., 2017). These changes progressively compromise cartilage mechanical integrity (Nimeskern L.U., 2015; Riedler K.L., 2017).

2.5 Tissue engineering and cartilage regeneration

The sparse distribution of differentiated chondrocytes, the slow turnover of matrix components and the low supply of progenitor cells cause an inadequate self-repairing ability of human cartilage (Ciorba A., 2006; Yamaoka H., 2006). The inability of self-regeneration and the shortcomings associated to surgical treatment of diseases that can cause cartilage deficiency, described in section 2.1, encourage scientists to investigate an efficient tissue engineering technique to regenerate cartilage in vitro. Tissue engineering is a multidisciplinary field which applies methods and principles of engineering and life sciences in order to produce biological substitutes able to ripristinate, mainatin or improve tissue functionality (Fuchs J.R., 2001). Three possible approaches have been investigated in order to regenerate functional tissue: cell ex vivo espansion before implantation in patients; tissue or organ ex vivo regeneration before implantation; design of devices able to stimulate tissue in vivo regeneration by inducing stem cells differentiation (Robey P.G., 2013). The success of tissue engineering techniques relies on three main elements: a safe and easily accessible cell source, a tunable culture environment and a three-dimensional biocompatible scaffold (Shieh S., 2004).

In 1997 Cao et al generated the first ear-shaped cartilage construct in an animal model (Cao Y., 1997), thus disclosing tissue engineering potential in auricle reconstruction. Since then many researchers have focused their attention on the possibility of regenerating an engineered human auricle to be implanted in patients who suffer of craniofacial development disorders. The first successful results were obtained by employing a two-steps scaffold-free approach: in the first step, chondrocytes isolated from microtic tissue were injected in an animal at subcutaneous level; successively the regenerated cartilage was harvested, manipulated and re-implanted in patients (Kamil S.H., 2004; Shieh S., 2004). Scaffold-based approaches were adopted in order to guarantee better mechanical support of cells during regeneration process. A polymeric scaffold

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16 supports development and maintenance of cartilage in a human ear shape after implantation (Isogai N., 2005; Kusuhara H., 2008). Vunjak-Novakovic et al suggested that cell culture within bioreactors, able to mimic dynamic physiological conditions experienced by cells, can improve engineered constructs mechanical properties and GAG synthesis (Vunjak-Novakovic G., 1999). Currently the production of engineered constructs having mechanical properties comparable with that of native elastic cartilage is still challenging. As underlined by Nimeskern et al, the elastic modulus of engineered cartilage samples reported in literature is < 1Mpa (Nimeskern L., 2015). The main consequence of poor mechanical strength is engineered cartilage difficulty to retain its complex three-dimensional shape after implantation (Ciorba A., 2006).

Recently one of the most investigated option is the use of additive manufacturing technologies to fabricate patient-specific scaffolds. Geometrical data are acquired by performing a CT scan of the healthy ear of the patient. These data are later processed to obtain a digital model and consequently a CAD negative model. This negative model is produced by additive manufacturing technologies and used as mould for the casting of the polymeric scaffold material, thus obtaining a bioscaffold that symmetrically replicate the three-dimensional structure of human auricle (Liu Y., 2010; Jessop Z.M., 2016; Zhou G., 2018). In wider terms this combination of CT scan acquisition and additive manufacturing technologies can be implemented to produce patient-specific implants and scaffolds that accurately replicate complex anatomical structures of the human body (Zopf D.A., 2018). Future perspectives aim at directly fabricating ear-shaped cartilage by using 3D bioprinting technique (Jessop Z.M., 2016). 3D bioprinting appears as a promising technology that allows to generate three-dimensional engineered constructs in complex shapes by bioink and cells deposition (Yang J., 2017). A study by Markstedt et al reported the fabrication of 3D bioprinted cartilage tissue by encapsulating chondrocytes in alginate-based hydrogel employed as bioink (Markstedt K., 2015). Main shortcomings of 3D bioprinting are: the lack of adequate mechanical properties and structural integrity; the low resolution that limit the control on micro and nanoscale architecture and finishing; the long processing time and shear stresses compromise encapsulated cells viability (Yang J., 2017). Hybrid bioprinting by combining a stiff synthetic polymer and a natural hydrogel is regarded as a feasible strategy to produce constructs characterised by high cell viability and improved mechanical stability (Park J.Y., 2017).

2.6 Cell source for in vitro chondrogenesis

Theoretically many cell types can be employed as cell source in cartilage engineering such as hMSCs, autologous chondrocytes, progenitor cells isolated from perichondrium, induced pluripotent stem cells (iPSCs) (Shieh S., 2004; Ciorba A., 2006; Makris E.A., 2015). Benefits and drawbacks of these cell sources are reported below.

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2.6.1 Mesenchymal stromal cells (MSCs)

MSCs were originally isolated from bone marrow by Friedenstein et al in 1966 as an adherent cell population similar to fibroblasts (Friedenstein A.J., 1966). MSCs include an heterogeneous population of multipotent cells including osteoprogenitors, adipocyte precursors and chondrogenic precursors, that can be isolated and cultured ex vivo for clinical use (Aubin J.E., 1995; Bernardo M.E., 2016). Indeed MSCs have the capability to extensively replicate and differentiate in vitro or in vivo in various mesenchymal tissues including bone, adipose tissue, muscle, tendon, cartilage (Pittenger M.F., 1999; Ciorba A., 2006; Jessop Z.M., 2016).

Figure 1 - Schematic illustration of MSC differentiation stages. Adapted from Aubin et al, 1995.

MSCs also exert a therapeutic effect by releasing a broad variety of bioactive molecules (trophic and regulatory proteins, including growth factors, proteinases, hormones, cytokines and chemokines), known as ‘MSC secretome’, able to contrast the inflammation and to support the regeneration of damaged tissue in vivo (Morigi M., 2016). Previous studies have already proved that bone marrow derived MSCs are able to differentiate in chondrocytes and to express chondrogenic markers, such as aggrecan and type II collagen when adequately stimulated (Ciuffreda M.C., 2016). The employment of MSCs in in vitro cartilage engineering presents several benefits as a low number of cells is initially required and their isolation from bone marrow is relatively easy and already adopted in clinical procedures (Ciorba A., 2006). However, prolonged in vitro culture is necessary in order to obtain noticeable chondrogenic differentiation (Liu Y., 2010). Moreover MSCs are not involved in the physiological development of auricolar cartilage and thus they are not suitable to produce elastic fibers present in the native tissue (Kusuhara H., 2009; Otto I.A., 2018).

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2.6.2 Differentiated chondrocytes

Studies conducted in 1990s demonstrated that both bovine chondrocytes and human chondrocytes are able to replicate and generate cartilage matrix when seeded on synthetic scaffolds (Vacanti C.A., 1994; Rodriguez A., 1997). However chondrocytes isolated from articular, costal or septal cartilage are not able to syntesize elastin, thus regenerated tissues show fibrocartilaginous nature and present lower biochemical and mechanical properties than native elastic cartilage (Jessop Z.M., 2016, Otto I.A., 2018). Moreover the use of microtic chondrocytes has been investigated as a potential cell source as they are able to produce elastic cartilage and they can be isolated from microtic ear without damaging healthy tissues (Ishak M.F., 2015). Nevertheless only a small amount of tissue can be harvested from the patient, thus an extensive in vitro expansion is required in order to obtain a sufficient number of cells to produce cartilage engineered constructs of clinically relevant size (Otto I.A., 2018). Furthermore chondrocytes proliferative potential is physiologically low and their capability to produce cartilage matrix decline over time (Otto I.A., 2018). By increasing the culture period and the number of passages chondrocytes progressively de-differentiate and lose their chondrogenic phenotype (Lee H.J., 2006; Yamaoka H., 2006). As a consequence of the de-differentiation chondrocytes assume a fibroblast-like morphology and there is a switch between type II and type I collagen synthesis (Otto I.A., 2018).

2.6.3 Cartilage stem/progenitor cells (CSPCs)

Kobayashi et al suggested the existence of a promising population of stem/progenitor cells that can be isolated from auricular perichondrium and guided to differentiate in chondrocytes (Kobayashi S., 2011). These cells, identified as cartilage stem/progenitor cells (CSPCs), are able to proliferate, differentiate and participate in tissue reconstruction in vivo like other adult stem cells, but they are characterised by a higher proliferative potential (Otto I.A., 2018). Moreover, as tissue-specific progenitor cells, CSPCs are considered more directed to differentiate in chondrogenic sense and exhibit an upregulated expression of elastin (Otto I.A., 2018). Though CSPCs proliferation rate is comparable with that of MSCs derived from bone marrow, MSCs show a more pronounced GAG deposition (Otto I.A., 2018). CSPC lower yield and the limited amount of generated matrix still restrict their use in experimental studies.

2.6.4 Human induced pluripotent stem cells (HiPSCs)

Autologous human induced pluripotent stem cells (HiPSCs), generated from adipose-derived stem cells (ASCs) or fibroblasts by viral reprogramming techniques, are studied as potential cell source in the regeneration of patient-specific cartilage (Nejadnik H., 2015). HiPSCs show a capability to express chondrogenic markers comparable or even superior to bone-marrow derived MSCs (Makris E.A., 2015). Moreover, HiPSCs may overcome some chondrocytes and MSCs shortcomings, such as the limited number

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19 of cells available from the donor, the invasiveness of harvesting procedure and the tendency of these cells to produce fibrous cartilage (Nejadnik H., 2015). Nevertheless there are still some unanswered questions related to the employment of iPSCs, such as their chondrogenic efficacy, an optimal protocol for isolation and differentiation ex vivo, undesired genomic alterations due to the viral reprogramming and iPSCs teratogenic potential in vivo (Makris E.A., 2015). The complex and inefficient process of HiPSC differentiation, in addition to the aforementioned shortcomings, limits the clinical applicability of this approach (Nejadnik H., 2015).

2.7 Chondrogenic culture conditions

Understanding the physiological mode of operation of chondrogenesis and what factors are able to influence this process is necessary in order to regenerate cartilage in vitro. Chondrogenesis is a multi-step process: undifferentiated MSCs aggregate and then differentiate in chondrocytes, successively these cells proliferate and synthesize matrix components (Akiyama, 2008). In the process of in vitro chondrogenesis, also defined as ‘artificial mesenchymal condensation’, cells adhere on the scaffold by forming aggregates in the attempt to recapitulate the embryonic process of ‘mesenchymal condensation’ that occurs in physiological chondrogenesis (Akiyama, 2008). Despite the broad number of studies dealing with hMSC in vitro chondrogenesis in pellet conditions dimensions, geometry and mechanical properties of the construct appears to be not physiologically relevant (Ng J., 2016; Pustlauk W., 2017). Moreover, hMSCs seeding on a scaffold can improve their capability to produce cartilage, by providing an environment that simulates the physiological one and by providing stimuli to facilitate cartilage development (Pustlauk W., 2017).

2.7.1 Cell seeding density influence on chondrogenesis

A key factor in processes of tissue regeneration in vitro is cell seeding density. An optimal density is required to guarantee the generation of appropriate cell-cell contacts able to induce cell differentiation and subsequent matrix deposition (Ingber D.E., 2006; Pustlauk W., 2017). Consequently, changes in seeding density of porous scaffolds may represent an efficient tool to control chondrogenic differentiation in vitro (Solchaga L.A., 2011; Pustlauk W., 2017). Huang et al asserted that chondrogenesis is limited by high seeding density due to negative feedback mechanisms and reduction of available nutrients and growth factors (Huang A.H., 2009).

2.7.2 Chondrogenic differentiative medium

The addition of proteins and growth factors to culture medium allows to mimic the physiologic microenvironment that improves cell proliferation and differentiation in vivo (Martin I., 2001). Indeed, cell interactions with ions, proteins and other molecules present in the culture medium can affect cell differentiative capability and consequently the quality and the amount of ECM produced by cells

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20 (Woodfield T.B.F., 2006). Several in vitro studies demonstrated that hMSCs are able to survive and differentiate in chondrocytes in the presence of a suitable culture medium consisting of Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with linoleic acid, serum albumin, insulin-transferrin-selenium (ITS), dexamethasone, transforming growth factor-beta 1 (TGF-β1), ascorbic acid, pyruvate sodium and Penicillin-Streptomicin (Pittenger M.F., 1999; Hansson A., 2017). Dexamethasone is an adrenocortical hormone capable to promote chondrogenic and osteogenic differentiation of stem cells and the synthesis of proteins typical of cartilage matrix in the presence of TGF-β (Yang J., 2017). TGF-βs identify a family of polypeptides regarded as a critical factor to both the in vivo and the in vitro induction of chondrogenesis, by facilitating initial cell adhesion and condensation (van Osch G.J., 1998). During chondrogenic differentiation TGF-β preserves chondrocytic phenotype and improves biochemical composition and functional properties of regenerated tissue, by regulating the expression of matrix components and the synthesis of degradative enzyme inhibitors (Makris E.A., 2015). The ascorbic acid plays a key role in the development of bone and cartilage tissues. Deficiency of ascorbic acid cause changes in collagen metabolism, thus reducing chondrocyte proliferation and matrix synthesis (Temu T.M., 2010). However a high concentration of ascorbic acid is cytotoxic (Choi K.M., 2008). Linoleic acid promotes cell differentiation by modulating cell metabolism and decreasing the proliferation rate (Hurley M.S., 2006). Insulin-transferrin-selenium (ITS) is a compound of insulin, transferrin and sodium selenium employed as commercial supplement within chondrogenic differentiative media, since it reduces the risk of chondrocytes hypertrophy and promotes the synthesis of aggrecan and type II collagen (Liu X., 2014).

2.8 Biomaterials and scaffolds

Biomaterials are employed to produce a cytocompatible, biocompatible, haemocompatible and sterilizable scaffold for tissue engineering (Ciorba A., 2006; Sterodimas A., 2009; Liu Y., 2010; Mallick K.K., 2013). The scaffold should be able to recapitulate native ECM characteristics and functions, guiding cell organisation, proliferation and differentiation in vitro by providing physical and biochemical cues (Sterodimas A., 2009; Mallick K.K., 2013). The ideal scaffold should be easily processed in a pre-defined shape and have adequate mechanical properties and stability to retain its shape during tissue regeneration process (Ciorba A., 2006; Sterodimas A., 2009; Liu Y., 2010; Mallick K.K., 2013). The biomaterial must have mechanical properties similar to that of the tissue to replace and of the surrounding healthy tissues in order to prevent mechanical mismatch and implant deformation (Griffin M.F., 2016). Moreover cells are sensitive to the mechanical properties of their microenvironment, therefore scaffold stiffness affect cell behaviour and fate (Bos E.J., 2017). Scaffold porosity and the degree of pore interconnectivity are critical in scaffold architecture. Interconnectivity and high porosity are necessary to guarantee an adequate infiltration of cells and nutrients within the scaffold, thus preventing the necrosis of cells located in the most internal region of the scaffold (Mallick K.K., 2013). However increased porosity reduces scaffold mechanical

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21 strength. As underlined in section 2.10, the capability to control pore interconnectivity and dimensions varies according to the fabrication technology adopted.

Several materials have been investigated as scaffold biomaterials for cartilage regeneration such as hyaluronic acid (HA) and autologous fibrin glue; poly(ethylene oxide); platelet-rich plasma; silicone; polytetraflorethylene (or Gore-tex); high-density porous polyethylene (HDPP or Medpor); poly(lactic acid) (PLA) and poly(glycolic acid) (PGA); poly(vinyl alcohol) (Cascone M.G., 2004; Moscato S., 2008; Griffin M.F., 2016; Liu Y., 2017). The ideal biomaterial has not been identified yet. Silicone leads to the formation of a fibrotic capsule around the implant with consequent poor attachment to the body; Gore-tex implants tend to loose their shape over time; Medpor has an elastic modulus two orders of magnitude higher than native auricular cartilage thus determining to an unnatural effect (Griffin M.F., 2016). Among the aforementioned materials, hydrogels, PLA/PGA and PVA are the most attractive.

2.8.1 Hydrogels

Among polymeric scaffolds, hydrogels appear to be particularly interesting due to the high hydration rate and the soft rubbery consistency make these materials similar to biological tissues (Drury J.L., 2003). Hydrogels consist of hydrophilic polymer chains linked together by chemical bonds or physical interactions that guarantee material structural integrity (Drury J.L., 2003). Despite their poor mechanical strength, hydrogels are more efficient in preserving the chondrocytic phenotype and supporting matrix synthesis (Yamaoka H., 2006). Currently hydrogels are widely investigated as printable biomaterials in the field of bioprinting (Jessop Z.M., 2016).

2.8.2 Poly(lactic acid) (PLA) and poly(glycolic acid) (PGA)

Poly(lactic acid) (PLA) and poly(glycolic acid) (PGA) are synthetic polymers frequently employed in tissue engineering scaffolds thanks to their tunable mechanical properties and the ease of shape processing (Liu Y., 2017). However these materials cause a severe foreign body reaction thus inhibiting cartilage formation in subcutaneous or intramuscular implant sites (Liu Y., 2017; Zhou G., 2018). In order to obviate this problem Liu et al suggested an in vitro pre-implantation culture to allow the formation of a mature cartilage tissue and adequate degradation of scaffold material before implantation (Liu Y., 2017).

2.8.3 Poly(vinyl alcohol) (PVA)

Poly(vinyl alcohol) (PVA) was one of the first synthetic polymers investigated to produce spongy-like matrix for tissue engineering and it is still one of the most attractive biomedical polymer thanks to its properties: biocompatibility, lack of toxicity, availability, low cost, FDA approval for use in biomedical applications (Moscato S., 2008; Kamoun E.A., 2015; Lee J.M., 2017). PVA can be physically crosslinked by freeze-drying or repeated cycles of freeze-thawing and chemically crosslinked with glutaraldehyde (GTA) (Drury J.L.,

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22 2003). Thanks to its hydrophilic nature, PVA is able to swell when immersed in an aqueous environment. Pure PVA shows limited poor capability to adsorb cells (Peng Z., 2011). In order to facilitate cell adhesion and resulting colonisation of the scaffold, PVA is frequently combined with biological molecules (Moscato S., 2008). In particular, gelatin/PVA bioscaffolds have been investigated and employed in tissue engineering applications (Cascone M.G., 2004; Kamoun E.A., 2015). Gelatin is a natural protein obtained from collagen denaturation. As the denaturation process preserves many functional groups of native collagen, gelatin is able to interact with cells and facilitate cell adhesion (Awad H.A., 2004). The most critical aspect to be considered in the employment of PVA-based sponges in tissue regeneration and replacement is the lack of adequate mechanical strength (Karimi A., 2014). Tensile mechanical tests performed on samples of PVA at different concentrations demonstrated that increasing polymer concentration in the aqueous solution significantly increases material elastic modulus (Schmedlen R.H., 2002).

2.9 Scaffold fabrication technologies

According to scientific literature, scaffold fabrication technologies can be divided in two main categories: conventional techniques (salt leaching, gas foaming, phase separation, free-drying) and solid free form fabrication techniques (3D printing, selective laser sintering, stereolithography, fused deposition modeling). Conventional techniques allow the production of spongy-like porous scaffolds, but pore interconnectivity and dimensions are difficult to control due to non-uniform dispersion of the porogen within the polymeric phase (Hutmacher, 2001). In solid free form and rapid prototyping technologies a three-dimensional scaffold is created by depositing and processing thin layers of materials starting from a computer-aided-design (CAD) model of the object (Hutmacher, 2001). Compared to conventional techniques, AM technologies allow the design and manufacture of fully interconnected porous scaffolds with a regular and reproducible morphology and eventually a variable microstructure (Hutmacher 2001; Loh Q.L., 2013). However a limited number of biomaterials can be processed by rapid prototyping techniques (Loh Q.L., 2013). Another easy and cheap fabrication technology is electrospinning. Electrospun scaffolds present high surface area and high porosity, but in solution electrospinning the use of organic solvents, harmful to cells, is required (Loh Q.L., 2013). As an alternative to scaffold prefabrication and successive cell seeding, cells can be encapsulated during scaffold fabrication process. Cells are mixed to the encapsulation material before gelification in order to trap the cells inside the material and protect them from organism immune response (Loh Q.L., 2013). Thus the material and the gelification process have to be biocompatible and non toxic. A balance between retention of immunoprotection and an adequate oxygen transport coefficient is necessary in order to improve the viability of encapsulated cells (Loh Q.L., 2013).

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2.10 Low-intensity ultrasounds effects on chondrogenesis

Static culture conditions are not sufficient to promote an adequate maturation of the engineered tissue. In addition to specific growth factors, mechanical stimuli such as cyclic loading are necessary to improve the maturation and functionality of engineered cartilage (Makris E.A., 2015). A deformational loading applied on the cell/scaffold construct improves cartilage in vitro regeneration by modulating ECM components synthesis (Lee H.J., 2006; Korstjens C.M., 2008; Ng J., 2016; Jonnalagadda U.S., 2018). In 2006 Lee et al investigated the effects of low-intensity ultrasounds treatment on the expression of chondrogenic markers and genes involved in matrix degradation during the chondrogenic differentiation of rabbit MSCs (Lee H.J., 2006). US stimulation results in a significant increase of cell proliferation and GAG, proteoglycan (in particular aggrecan) and type II collagen synthesis, while the expression of type I collagen significantly decreases (Parvizi J., 1999; Nishikori T., 2002; Xia P., 2017; Aliabouzar M., 2018; Jonnalagadda U.S., 2018). These results can be regarded as a proof of cellular chondrogenic differentiation. Moreover, it has been observed that the effect of LIUS combined with the presence of dexamethasone and TGF-β1 in the culture medium leads to an increase in mRNA expression of chondrogenic markers such as sex determining region box 9 (Sox9) (Korstjens C.M., 2008; Lai C.H., 2010). Sox9 can be identified as ‘the master regulator’ of chondrogenesis, indeed it is a transcription factor necessary to mesenchymal condensation, proliferation and differentiation (Akiyama, 2008). In particular it has been demonstrated that the expression of Sox9 induces cell expression of type II collagen, typical of the cartilagineous matrix (Ikeda T., 2004). A fundamental factor to be considered is the intensity of LIUS: low intensity does not cause a mechanical stimulation sufficient to induce cell proliferation; while high intensity can cause excessive shear stress values and damage cell membrane (Aliabouzar M., 2018). A 200 mW/cm² US intensity is regarded as an optimal value to promote in vitro chondrogenesis (Lee H.J., 2006; Lai C.H., 2010). The mechanism by which LIUS can affect chondrogenesis is not well understand but probably includes mechanotrasduction pathways mediated by integrins (Lai C.H., 2010).

2.11 Aim of the study

The overall aim of the project is the in vitro regeneration of auricular cartilage by differentiating hMSCs, seeded on a bioartificial ear-shaped scaffold. More particularly, the aim of this study is the fabrication and characterisation of a bioartificial scaffold for auricular cartilage engineering. Poly(vinyl alcohol) (PVA), a biocompatible and polymeric synthetic polymer was combined with gelatin and alginate to produce hydrophilic sponges able to emulate native extracellular matrix. Human mesenchymal stromal cells (hMSCs) were employed as cell source since they can be easily isolated from bone marrow and they are able to extensively replicate in vitro and differentiate in chondrocytes. Low-intensity ultrasound (LIUS) stimulation was considered, since previous studies demonstrated LIUS ability to induce an increase in

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24 chondrogenic markers expression. Specific objectives of the study are the identification of the optimal scaffold composition and the assessment of an optimal hMSC differentiation protocol.

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25

Chapter 3 Materials and Methods

3.1 Scaffold fabrication

PVA-based scaffolds were produced via emulsion and freeze-drying (Moscato S., 2008; Barachini S., 2014). A 11.7% aqueous solution of PVA (Mw=89000-98000 g/mol, >99% hydrolysed from Sigma-Aldrich, St. Louis, MO, USA) in Milli-Q® deionized water was autoclaved 1 h at 120°C and then was cooled down to 50°C inside a thermostatic bath under stirring at 1000 rpm. Gelatin (gelatin from bovine skin, type B, from Sigma-Aldrich) and Alginic acid sodium salt (Fluka BioChemika) were added in order to obtain different polymer/biomolecule composition: PVA/G 90/10, 80/20, 70/30 and PVA/G/alginate 80/10/10 and 90/5/5 (w/w)%. The temperature was increased to 70°C to allow an optimal biomolecule dissolution and then decreased again to 50°C always under stirring. 0.18 gr of sodium dodecyl sulfate (SDS, from Sigma-Aldrich) were added to obtain a dense foam. After 10 minutes under stirring the foam was poured into a six-well plate, quenched in liquid nitrogen and lyophilized. The result of this process is a spongy-like highly porous and hydrophilic scaffold. Dried foams were crosslinked by exposure to glutaraldehyde (GTA, grade II, from Sigma-Aldrich) vapours for 72 h at 37°C in a sealed cabinet. After crosslinking, sponges were flushed under the chemical hood for 72 h.

3.2 Scaffold characterisation

3.2.1 SEM analysis

PVA/G samples (90/10, 80/20 and 70/30 w/w%) were mounted on aluminium stumps, sputter-coated with gold (Sputter Coater Emitech K550, Quorum Technologies Ltd, West Sussex, United Kingdom) and examined on a scanning electron microscope (JEOL JSM-5200, JEOL Ltd, Tokyo, Japan).

3.2.2 Swelling analysis

5 cylindrical samples of each composition were obtained by using a 5 mm diameter biopsy puncher. Dry sample dimensions were measured using a digital caliper (resolution=0.1 mm). Dimension measurements were repeated at different time point on wet samples. Volume swelling ratio (Q, Equation 1) was calculated according to the following equation (Karimi A., 2014):

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26

3.2.3 Gelatin release

Gelatin release from crosslinked and uncrosslinked PVA/G sponges was evaluated by means of a spectrophotometric method. 5 mg of different (w/w)% composition PVA/G sponges were immersed in 1 ml of phosphate buffer saline (PBS) and incubated at 37°C for 96 h. Then, release solution was collected from each sample and diluted 1:10 in 1x PBS. 200 μL of Bradford reagent (Bio-Rad, USA) were added to 800 μL of test solution. The absorbance of the solution at 595 nm was measured using a photometer (BioPhotometer plus, Eppendorf, Germany). Gelatin concentration in the release solution was determined through comparison with a calibration curve.

3.2.4 Differential scanning calorimetry (DSC)

Phase transition properties of the samples were determined using a DSC Q200 Differential Scanning Calorimeter controlled by a TA module (TA Instruments, New Castle, USA). 7-8 mg of dry samples were placed in hermetically sealed aluminum pans and heated from -35 to 250°C at a heating rate of 10°C/min in an inert atmosphere. An empty pan was used as the reference cell. Transition temperatures were calculated using the Universal Analysis 2000 software (TA Instruments).

3.2.5 Fourier transformed infrared spectroscopy (FTIR)

FTIR spectra were recorded using Nicolet 380 FT-IR Spectrometer (ThermoFisher Scientific, USA) equipped with the Thermo Scientific™ Smart™ iTX accessory. The spectra were recorded between 4000 and 550 cm¯¹ with a 8 cm¯¹ spectral resolution. For each spectrum 256 scans were co-added. The FTIR spectrum was taken in a transmittance mode. Data were analysezed using EZ OMNIC Software (ThermoFisher Scientific, USA).

3.2.6 Mechanical testing

Viscoelastic properties of PVA/G sponges at different (w/w)% ratios were investigated by using epsilon dot method (Mattei G., 2014; Tirella A., 2014). Mechanical tests were performed in triplicate. Samples were equilibrium swollen in distilled water at room temperature. Specimens dimensions were measured with a digital caliper after swelling. Then, samples were fixed at the bottom of Petri dishes and some drops of distilled water were placed around them to prevent dehydration throughout testing. Short compressive test were performed at different strain rates (0.01, 0.005, 0.001, 0.0005 s¯¹) using the twin column ProLine Z005 testing machine (Zwick Roell) equipped with a 10 N load cell (Zwick KAP-TC). In the start configuration the upper plate is near the sample but not in contact to guarantee a zero stress initial condition. Test was controlled in position. Force and displacement data were recorded over time (time sampling 1 ms). Firstly

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27 data were processed in Excel to obtain stress-time series. Stress (Equation 2) and strain (Equation 3) values were calculated as follows:

(Equation 2)

where F is the force measured by the load cell and A₀ is sample cross-sectional area; (Equation 3)

where Δl is the displacement recorded and l₀ is sample initial thickness. Apparent elastic moduli were calculated as the slopes of stress-strain curves and linear viscoelastic region was identified. A standard linear solid (SLS) model was chosen to derive PVA/G viscoelastic parameters. Equation 4 represents the constitutive equation of a Maxwell-type SLS model.

( ) ̇ ( _{) (Equation 4)}

Viscoelastic parameters ( ) were estimated by globally fitting stress-time series in a combined parameter space using OriginLab (Northampton) fitting toolbox (Mattei G., 2014; Tirella A., 2014). Instantantaneous elastic modulus (Einst), equilibrium elastic modulus (Eeq) and relaxation time (τ) were derived according to the following equations (Equation 5, 6, 7).

(Equation 5) (Equation 6)

(Equation 7)

3.3 Biological studies

PVA/G 70/30 (w/w)% sponges were cut in cylindrical scaffolds (5 mm in diameter and 1.5 mm in thickness) using a biopsy puncher and a microtome blade. Scaffolds were sterilised in absolute ethanol (Bio-optica, Milan, Italy) for 24 hours and then treated with 2% glycine (Sigma-Aldrich) for 1 hour in order to block unreacted sites of GTA. Therefore, scaffolds were washed three times in phosphate buffered saline (PBS) supplemented with antibiotics and then moved into sterile multiwell plate.

3.3.1 hMSC isolation from bone marrow

Human mesenchymal stromal cells (hMSCs) were isolated from bone marrow aspirates according to Ficoll protocol (Seeger F.H. 2007). Bone marrow asprirates were obtained from adult patients undergoing routine total hip replacement surgery, after written consent. 20 ml of bone marrow aspirate were diluted 1:4 with

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28 0.9% NaCl and mononuclear cells were isolated by density gradient centrifugation at 1500 rpm for 15 minutes without brake using Ficoll. Mononuclear cells were then collected and centrifugated again at 1500 rpm for 5 minutes. Supernatant was discarded and cells were resuspended in Dulbecco’s Modified Eagle Medium Low Glucose (DMEM-LG) supplemented with 10% fetal bovine serum (FBS), 100 IU/ml penicillin (Pharmacia & Upjohn S.p.A., Milan, Italy), 100 μg/ml streptomycin (Bristol-Myers Squibb S.p.A., Sermoneta, Italy), 2mM L-glutamine (Lonza, Wolkersville, USA), 2 mg/ml fluconazole. Cells were seeded at a density of 250 000 cells/cm² and cultured in standard cell culture conditions (37°C, humidity>95%, 5% CO₂). After 48-72 hours nonadherent cells were discarded and fresh medium was added. When 80% confluence was reached adherent cells were trypsinized and seeded at low density for further expansion (Trombi L., 2008; Penfornis P., 2016).

3.3.2 Chondrogenic differentiative medium

An homemade chondrogenic medium was obtained starting from DMEM/F12 by adding 1.25 μg/ml bovine serum albumin (BSA), 10 μl/ml insulin-transferrin-selenium (ITS premix), 5.35 μg/ml linoleic acid, 50 μg/ml ascorbic acid, 100 μg/ml pyruvate sodium, 10¯⁷M dexamethasone, 10 ng/ml transforming growth factor beta 1 (TGF-β1, PeproTech, Rocky Hill, New Jersey) (Solchaga L.A., 2011; Barachini S., 2014; Ciuffreda M.C., 2016). If not otherwise specified, all reagents were purchased from Sigma-Aldrich.

3.3.3 Cell viability

AlamarBlue assay was performed once a week in order to assess cell viability. Cell/scaffold constructs to be tested were placed in 15 ml Falcon tubes and alamarBlue® reagent (Life Technologies, USA) was added, according to manufacturer’s prescriptions. AlamarBlue solution without cells served as negative controls. After 3 hours of incubation, 100 μl samples were collected in triplicate and analysed with a plate reader (Victor3, PerkinElmer, Waltham, USA). Absorbance values at 570 nm and 600 nm were recorded and used to calculate dye reduction percentage. At the end of the test fresh culture medium was added.

3.3.4 Low intensity ultrasounds stimulation

Cells were stimulated with low intensity ultrasounds at 40 kHz frequency and 20 W output power using a sonicator bath (Bransonic 2510; Bransonic, Danbury, USA) as reported by Barachini et al. (Barachini S., 2014). A set of cell/scaffold constructs were daily treated 3 times for 5 seconds. Non-stimulated constructs served as negative controls. US treatment was started after 1 day of culture in differentiative medium.

3.3.5 First experiment: 14 days differentiation

A preliminary experimental study was performed in order to evaluate scaffold ability to support hMSC growth and chondrogenic differentiation in vitro. Twice passaged hMSCs were seeded on PVA/G 70/30

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29 scaffolds at a seeding density of 500 000 cell/scaffold (Barachini S., 2014) and cultured in StemMACS ChondroDiff (Miltenyi Biotec, Germany) differentiative medium for 14 days. Medium changes were carried out twice a week. At the endpoint samples were processed for scanning electron microscopy and histological/immunohistochemical analysis.

3.3.6 Second experiment: 21 days differentiation

The second experimental study was designed to identify the best differentiative conditions as detailed below. In a first step hMSCs at passage 2 were trypsinized and seeded on PVA/G 70/30 scaffolds (5 mm in diameter and 1.5 mm in thickness) at a density of 500 000 cell/sample (n=6). After 1 day the culture medium was replaced with chondrogenic differentiative medium. Milteniy ChondroDiff medium was added to a set of samples (n=3) and a homemade chondrogenic medium, prepared as reported in section 3.3.2, was added to the other set (n=3). Cells were cultured in standard conditions and differentiated for 21 days. Two samples of each group were daily treated with LIUS as described in section 3.3.4. Non-US stimulated samples served as negative controls. In a second step hMSCs at passage 2 were trypsinized, placed in 15 ml tubes at a density of 250 000 cells/tube and centrifuged at 1200 rpm for 7 min to obtain chondrogenic pellets as described by Barachini et al (Barachini S., 2014). Pellets were cultured 1 week in Milteniy ChondroDiff medium, then trypsinized, seeded on PVA/G 70/30 scaffolds (5 mm in diameter and 1.5 mm in thickness) and cultured for 14 days. Cell/scaffold constructs were daily treated with LIUS as described in section 3.3.4. Finally, hMSCs at passage 2 were trypsinized and seeded in 25 cm² flasks at a seeding density of 5 000 cells/cm² (Bajpai V.K., 2012). After 4 days of expansion in DMEM-LG, cells were committed to chondrogenic lineage by adding Milteniy ChondroDiff medium and pre-differentiated for 4 days. After the pre-differentiation in 2D conditions, cells were trypsinized, seeded on PVA/G 70/30 scaffolds (5 mm in diameter and 1.5 mm in thickness) at a density of 300 000 cell/sample (n=2) and differentiated for 17 days. Samples were daily treated with LIUS as described in section 3.3.4. AlamarBlue assays were performed at different time points to assess cell viability throughout culture time. Medium changes were carried out twice a week. At the endpoint all samples were processed for scanning electron microscopy and histological/immunohistochemical analysis.

3.3.3 Third experiment: 24 days differentiation

Twice passaged hMSCs, pre-differentiated in 2D conditions as decribed in the previous section, were trypsinized, seeded on PVA/G 70/30 scaffolds (5 mm in diameter and 1.5 mm in thickness) at a density of 300 000 cell/sample (n=8) and differentiated for 24 days. A set of samples (n=4) was daily treated with LIUS as described in section 3.3.4 extending stimulation time by 5 seconds, while the other set (n=4) served as negative control. AlamarBlue assays were performed to assess cell viability throughout culture time.

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30 Medium changes were carried out twice a week. At the endpoint all samples were processed for scanning electron microscopy and histological/immunohistochemical analysis.

3.4 Histological analysis

3.4.1 Histological sample preparation

Cell/scaffold constructs were fixed in 4% w/v neutral buffered formalin diluted in 1x PBS (0.1M pH 7.2) (Bio-Optica) overnight at 4°C. Thereafter samples were washed in 1x PBS and dehydrated with a graded series of ethanol (from 70% to 100%). After 3 h of incubation in absolute ethanol (Bio-Optica), samples were clarified in xylene (2 steps of 45 min each one) (Sigma-Aldrich). All steps were performed in a thermostatic bath at 40°C. Successively, specimens were rinsed in liquid paraffin (Histoplast LP, Thermo Fisher Scientific, Waltham, MA, USA) at 60°C for 2 h and paraffin-embedded. 8-micron-thick sections were obtained by standard microtome and mounted onto glass slides. Before each histological staining or immunoreaction, sections were deparaffined in xylene (2 steps of 7 min each one), rehydrated in absolute ethanol (3 steps of 7 min each one) and rinsed in distilled water for 5 min.

3.4.2 Hematoxylin & Eosin Staining

Hematoxylin stains cell nuclei in blue, Eosin stains cytoplasm in pink.

Deparaffinized sections were stained with Mayer’s hematoxylin for 5 min, washed in tap water for 5 min, counterstained with eosin for 1 min and then washed in distilled water (all reagents were from Sigma-Aldrich). Samples were then dehydrated in absolute ethanol (3 steps of 5 min each one), clarified in xylene (3 steps of 5 min each one) and mounted in DPX medium (Sigma-Aldrich).

3.4.3 Periodic acid-Schiff (PAS) Staining

Periodic acid oxidizes 1,2 glycol groups to aldehyde groups. Schiff reagent reacts selectively with these aldehydes, thus staining glycoproteins in the extracellular matrix.

Specimens were oxidized in 1% w/v periodic acid (Sigma-Aldrich) solution for 10 min, air dried and then incubated in Schiff reagent (Carlo Erba) for 15 min. Then, sections were washed in tap water for 10 min, counterstained with Mayer’s hematoxylin for 5 min and finally rinsed in tap water for 5 minutes. Specimens were then dehydrated, clarified and mounted as previously described.

3.4.4 Alcian Blue Staining

Alcian blue pH 1 highlights the presence of sulfated acid proteoglycans, typical of cartilge matrix, while Alcian blue pH 2.5 shows generic acid proteoglycans.

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31 Specimens were incubated in Alcian Blue pH 1 and pH 2.5 solutions (Bio-Optica) for 30 min and in revealing solutions (Bio-Optica) for 10 minutes, according to manufacturer’s instructions. After the incubation time, sections were washed in distilled water, counterstained with a distilled water solution containing 0.1% w/v Nuclear Fast Red (Sigma-Aldrich) and 5% w/v Aluminium Sulfate (Carlo Erba, Milan, Italy) for 5 minutes and washed in tap water for 5 minutes. Specimens were finally dehydrated, clarified and mounted as previously described.

3.4.5 Toluidine Blue staining

Toluidine Blue is an acidophilic metachromatic dye that stains in violet acidic components, like sulfated acid glycosaminoglycans.

Sections were incubated with a distilled water solution containing 0.2% w/v Toluidine Blue (Sigma-Aldrich) and 1% w/v sodium tetraborate decahydrate, for 10 s. Then, sections were rinsed in tap water and air dried. Finally, samples were clarified in xylene and mounted in DPX medium.

3.4.6 Orcein-Van Gieson Staining

Orcein highlights the presence of elastic fibers in violet. Van Gieson stains collagen fibers in red.

Sections were incubated in an ethanolic solution containing 1% w/v orcein for 1 h and then rinsed in 1% HCl (Carlo Erba) acidified 70% ethanol (2 steps of 5 min each one). Then, sections were washed in tap water (2 steps of 5 min each one), counterstained in Mayer’s hematoxylin for 5 min and washed in tap water for 10 min. Specimens were rinsed in distilled water, incubated in Van Gieson solution containing 0.1% w/v acid fuchsin (Sigma-Aldrich) in saturated picric acid (Sigma-Aldrich) for 2 min, rinsed in distilled water and air dried. Finally, samples were clarified in xylene and mounted as previously described.

3.4.7 Alcian Blue pH 1-Van Gieson Staining

As previously described, Alcian blue pH 1 highlights the presence of sulfated acid GAGs and Van-Gieson stains collagen fibers in red.

Specimens were incubated in Alcian Blue pH 1 solution and in revealing solution (Bio-Optica), as described in section 3.4.3, then counterstained in Nuclear Fast Red for 5 min and washed in tap water for 3 min. Then, sections were incubated in Van Gieson solution for 2 min, rinsed in distilled water and air dried. Finally, samples were clarified in xylene and mounted in DPX medium.

3.5 Immunohistochemical analysis

Sections were washed in 1x PBS for 5 min. Sections employed for the detection of type I collagen and sox-9 were permeabilized with 0.2% w/v Triton X-100 (Sigma-Aldrich) in 1x PBS for 10 min. Sections employed for the detection of aggrecan, type II collagen and elastin were unmasked by incubation in citrate buffer